By the time Watson and Crick described the structure of DNA in 1953, its loss of structure at high pH or temperature was already well known. The double-helix structure made it obvious that DNA could be unzipped by breaking the hydrogen bonds holding the two helices together; this was the mechanism by which DNA melting occurred. Early work focused on synthetic DNA sequences found that the length of the DNA sequence and percentage of CG base pairing were directly proportional to increases in melting temperature (Tm) or pH. Today, in addition to its uses as a measure of DNA length and % CG content, DNA melting curves are used to elucidate epigenetic modifications (DNA methylation) and to detect single nucleotide polymorphisms (SNPs) in individual genes.
These advances have been possible due to adaptation of cellular DNA replication machinery to synthetic gene amplification by PCR. This technique uses fluorescent dyes to measure increases in DNA concentration during temperature cycling. Once amplification is complete, these dyes enable post-PCR melting curves to verify Tm as a test of amplification specificity. Because DNA melts within a narrow temperature range (a few ° C.), melting curve acquisition requires the imaging and temperature control capabilities provided by a real-time thermocycler.
Such DNA melting curves are possible because these dyes fluoresce preferentially when intercalated into double-stranded, but not single-stranded DNA. As the DNA denatures, the dye is no longer stabilized by the negatively charged DNA backbone and the fluorescence is quenched. This mechanism is common to most DNA-binding dyes; however the degree of double-stranded specificity is a property of each dye. If the melting curve is preceded by amplification, a large amount of specificity is not required as the amplification step reduces any nonspecific binding to levels below the noise threshold. However, if no amplification is performed, then this dye must be very specific for double-stranded DNA (dsDNA) (more than 100-fold over single-stranded DNA (ssDNA) and RNA).
To date, determination of genotoxicity is extremely low throughput. Many current assays measure either bacterial or mammalian cell growth in media treated with potential mutagens. Other cell-based assays take a gene induction-based approach, using reporter assays to measure induction of key DNA damage proteins in mammalian cell lines, such as repair enzymes. However the cellular machinery for fixing DNA double-strand breakage, single-strand breakage, and nucleotide base damage are vastly different, necessitating separate tests. Furthermore, there is cross talk between the pathways mediating signal induction, which can lead to false positive and false negative tests. Attempts to increase throughput of cell-based assays using microtiter plate readers have floundered because ensuring the quality of these assays between plates is difficult. Current in vitro genotoxicity tests, such as comet assays and 8-hydroxy-2-deoxyguanosine production, ignore some forms of DNA damage, like adduct formation and DNA strand cross linking, in favor of others. In the end, multiple assays must be performed and disparate data must be compared. None of these techniques readily lends itself to testing multiple compounds at the same time and the use of compression is vital when testing a library containing millions of compounds.
Three groups have used highly specific dsDNA binding dyes to quantify DNA damage by melting curve analysis (Singer et al. Anal Biochem. 1997 249:228-238; Kailasam et al. Chemosphere 2007 66:165-171; Rogers K R and Apostol A, Anal Chem 1999 71: 4423-4426; Batel et al., Anal. Biochem. 1999 270:195-200). The first measured changes in the rate of dsDNA denaturation at high pH and correlated this to radiation exposure. However this technique requires the entire DNA to be of the same sequence and length and the pH of the buffer to be exactly determined, making it impractical for use as a high-throughput assay. The second group used temperature to denature the radiation-damaged dsDNA. However, their methodology was flawed as they performed measurements after heating the DNA to only three temperatures in a waterbath and required the fluorescence of the DNA binding dye, in this case PicoGreen, be determined immediately, before the DNA could re-anneal. Again, this technique requires the DNA to be of the same sequence and length and does not lend itself to a high-throughput screen. The third group measured loss of fluorescence after successive melting and re-annealation qPCR cycles. This decrease in fluorescence is attributed to the loss of small DNA fragments resulting from radiation damage. While this endpoint assay is the most high-throughput of the three techniques, it still quantifies only DNA strand breakage since it does not measure the entire melting curve, but only the initial value of each cycle. Nor does it accurately quantify DNA strand breakage, since the large DNA fragments formed could still re-anneal (U.S. Pat. No. 4,407,942 10/1983 and U.S. Pat. No. 5,863,753 1/1999).
In light of the foregoing, it would significantly advance the art if a quantitative method for detecting genotoxicity was developed that could simultaneously differentiate between different forms of DNA damage formation. It would be an additional advancement if such an assay could take advantage of high-throughput technological developments such as fluidics automation and dense microtiter formats to produce rapid, accurate results without being labor-intensive. Such a method is disclosed herein.